Disturbed vitamin A metabolism in chronic liver disease and relevance for therapy
Saeed, Ali
IMPORTANT NOTE: You are advised to consult the publisher's version (publisher's PDF) if you wish to cite from
it. Please check the document version below.
Document Version
Publisher's PDF, also known as Version of record
Publication date:
2019
Link to publication in University of Groningen/UMCG research database
Citation for published version (APA):
Saeed, A. (2019). Disturbed vitamin A metabolism in chronic liver disease and relevance for therapy.
University of Groningen.
Copyright
Other than for strictly personal use, it is not permitted to download or to forward/distribute the text or part of it without the consent of the author(s) and/or copyright holder(s), unless the work is under an open content license (like Creative Commons).
Take-down policy
If you believe that this document breaches copyright please contact us providing details, and we will remove access to the work immediately and investigate your claim.
Downloaded from the University of Groningen/UMCG research database (Pure): http://www.rug.nl/research/portal. For technical reasons the number of authors shown on this cover page is limited to 10 maximum.
Chapter 5
Impaired
hepatic
vitamin
A
metabolism in NAFLD mice leading
to vitamin A accumulation in
hepatocytes
Ali Saeed
1,2, Paulina Bartuzi
3, Janette Heegsma
1,4, Daphne
Dekker
3, Niels Kloosterhuis
3, Alain de Bruin
3,5, Johan W.
Jonker
6, Bart van de Sluis
3, Klaas Nico Faber
1,4$1Department of Gastroenterology and Hepatology, 4Laboratory Medicine, Department
of Pediatrics, 3Section of Molecular Genetics and 4Section of Molecular Metabolism
and Nutrition, University Medical Center Groningen, University of Groningen, Groningen, The Netherlands. 5Dutch Molecular Pathology Center, Department of
Pathobiology, Faculty of Veterinary Medicine, Utrecht University, Utrecht, The Netherlands.
2Institute of Molecular Biology and Biotechnology, Bahauddin Zakariya University
Multan, Pakistan.
144
ABSTRACT
Vitamin A homeostasis is centrally controlled by the liver, involving close collaboration between hepatocytes and hepatic stellate cells (HSC). Non-alcoholic fatty liver disease (NAFLD), as well as other chronic liver diseases, is associated with low hepatic and serum retinol levels, suggestive of systemic vitamin A deficiency. In this study we show that, in contrast to retinol, hepatic levels of retinyl palmitate (the main vitamin A storage form) are strongly enhanced in 2 in vivo mouse models of NAFLD. Transcriptome analysis supports the simultaneous enhancement of retinol conversion to retinyl esters and retinoic acids in NAFLD livers. Vitamin A accumulates in hepatocytes in NAFLD, while this occurs in HSC in control mice, findings corroborated by palmitic acid-treated hepatocytes and HSC in vitro. Thus, NAFLD does not lead to true vitamin A deficiency, but to cell type-specific rearrangements in vitamin A metabolism leading to hepatic retinyl ester accumulation and retinol depletion.
145
5.1. INTRODUCTION
Non-alcoholic fatty liver disease (NAFLD) is the most common liver disease worldwide. The prevalence of NAFLD is estimated at 20-30% in the general population in Western countries. Starting with benign steatosis, patients are at risk to develop non-alcoholic steatohepatitis (NASH), progress to cirrhosis and hepatocellular carcinoma [1]. NALFD prevalence is particularly high in obese individuals (80-90%), diabetes (30-50%) and/or hyperlipidemia (90%) [1,2]. Hepatic fat accumulation is a combined result of a high fat- and high carbohydrate-containing diet and insufficient catabolism of these energy sources, in part due to low physical activity [3]. A tipping point NAFLD pathology is when simple steatosis is being accompanied by hepatic inflammation and fibrosis. Inflammation leads to the activation of hepatic stellate cells(HSC) that transdifferentiate to proliferative and migratory myofibroblasts, e.g. activated HSC, that produce excessive amounts of extracellular matrix proteins, like collagens and fibronectins, the typical feature of fibrosis [4].
HSC is considered “quiescent” in the healthy liver, though they play a key role in controlling vitamin A metabolism [5]. Vitamin A is important for many physiological processes, including reproduction, embryogenesis, glucose- and lipid metabolism and vision, most of which are controlled by the retinoic acid-activated transcription factors retinoid X receptor (RXR) and retinoic acid receptor (RAR). Quiescent HSC contains the main body reserve of vitamin A, which is stored as retinyl esters in large cytoplasmic lipid droplets [6,7]. Controlled conversion to retinol in qHSC is believed to maintain stable circulating serum retinol levels around 2.0 µmol/L in humans (1.0-1.5 µmol/L in rodents). The specific (control) mechanisms involved are, however, largely unknown. Dietary vitamin A follows a complex route for final storage in HSC [7]. In the small intestine, retinyl esters are incorporated in chylomicrons and after transport through the circulation taken up by hepatocytes. In hepatocytes, they are converted to retinol and subsequent binding to retinol binding protein 4 (RBP4) stimulates the release of the retinol-RBP4 complex back to the circulation. Via an unknown mechanism, retinol is transferred to HSC, re-esterified and stored in lipid droplets. Liver injury-induced activation of HSC leads to the rapid loss of vitamin A-containing lipid droplets and, as a consequence, chronic liver diseases are associated with vitamin A deficiency (VAD), including viral hepatitis, primary biliary cholangitis (PBC), primary sclerosing cholangitis (PSC), biliary atresia, alcoholic hepatitis and also
146
NAFLD [8]. VAD is defined as serum retinol levels below 0.7 mmol/L [9]. Not only serum, but also hepatic retinol levels were found to be reduced in class-III (BMI≥40 kg/m2) obese patients and negatively correlated with histological severity of hepatic
steatosis [10,11]. These analyses, however, do not take retinyl esters into account, which form the largest pool of vitamin A. It thus remains unclear whether the low serum and hepatic retinol levels truly reflect complete depletion of vitamin A stores or, alternatively, point to aberrant vitamin A metabolism in the fatty liver. Notably, individuals carrying the NAFLD risk variant of patatin-like phospholipase domain-containing protein 3 (PNPLA3, encoding adiponutrin) show low serum retinol levels, while hepatic retinyl ester levels are increased, pointing to impaired hepatic retinyl ester-to-retinol conversion [12]. Moreover, transcriptome analyses revealed a hyperdynamic state, of hepatic retinol metabolism in NAFLD patients, e.g. enhanced expression of genes involved in vitamin A storage, hydrolysis and transport, though with unknown effects on the various vitamin A pools [13].
Thus, in this study we analyzed serum and hepatic retinol, retinyl ester and RBP4 levels in 2 NAFLD mouse models, as well as expression of genes/proteins involved in vitamin A metabolism. Our data show that NAFLD in mice does not lead to plain vitamin A deficiency, but enhances vitamin A storage in the liver while retinol levels are reduced. Importantly, the bulk of the vitamin A accumulates in hepatocytes, rather than in HSC.
5.2. MATERIALS AND METHODS
Animals
All animal experiments were approved by the Institutional Animal Care and Use Committee, University of Groningen, The Netherlands. Age- and sex-matched (8-10 weeks old) C57BL/6J mice, (Charles River, Saint-Germain-Nuelles, France) were fed either regular chow or a high fat, high cholesterol (HFC) diet (containing 21% fat, with 45% calories from butter-fat, 0.2% cholesterol and per gram of diet) (Scientific Animal Food and Engineering (SAFE), Villemoisson-Sur-Orge, France) for period of 12 or 20 weeks (n=8). Both diets contained 20 IU of retinyl acetate/g as a source of vitamin A. Leptinob mutant (JAX™ ob/ob) mice (10-12 weeks; Charles River) on a C57BL/6J
genetic background and appropriate controls (C57BL/6J) were also analyzed and kept on chow diet. All animals were kept in a pathogen-free environment with
147
alternating dark-light cycles of 12 hours, and controlled temperature (20-24 ºC) and relative humidity (55%±15%). Animals received food and water ad libitum. All animals were fasted 4 hours before sacrifice. Tissues were snap-frozen in liquid nitrogen or fixed in paraformaldehyde. Blood was collected by heart puncture.
Primary hepatocyte increased the accumulation of vitamin A in high fat In vitro
Primary rat hepatocytes and hepatic stellate cells (HSC) were isolated from Wister rat (Charles River) and cultured in William’s E medium (Thermo Fisher Scientific, Breda, The Netherlands) and Iscove’s Modified Dulbecco’s Medium with Glutamax (ThermoFisher, Scientific), respectively, in a humidified incubator at 37ºC and 5% CO2, as previously described [14–16]. Primary hepatocytes and HSC were exposed
to BSA-conjugated palmitic acid (Sigma-Aldrich, St Louis, USA) (0.25 or 0.5 mmol/L), with or without retinol (Sigma-Aldrich) (5 µM) for 24-48 hours as previously described [17].
Quantitative real-time reverse transcription polymerase chain reaction (qRT-PCR)
Quantitative real-time reverse transcription polymerase chain reaction was performed as previously described [18]. Shortly, total RNA was isolated from tissue samples using TRIzol® reagent according to the supplier’s instructions (ThermoFisher
Scientific). The RNA quality and quantity were determined using a Nanodrop 2000c UV-vis spectrophotometer (Thermo Fisher Scientific). cDNA was synthesized from 2.5 µg of RNA by using random nonamers and M-MLV reverse transcriptase (Thermo Fisher Scientific). Taqman primers and probes were designed using Primer Express 3.0.1 (ThermoFisher, Scientific) and are shown in Supplementary Table
S1. All target genes were amplified using the Q-PCR core kit master mix
(Eurogentec, Maastricht, The Netherlands) on a 7900HT Fast Real-Time PCR system (ThermoFisher, Scientific). SDSV2.4.1 (ThermoFisher, Scientific) was used to analyze the data. Expression of genes is presented in 2-delta CT and normalized to 36B4.
148
Western blot analysis
Protein samples were prepared for Western blot analysis as described previously [19]. Protein concentrations were quantified using the Bio-Rad protein assay (Bio-Rad, Hercules, CA, USA) with bovine serum albumin (BSA) as standard. Equal amounts of protein were separated on Mini-PROTEAN® TGX™ precast 4-15%
gradient gels (Bio-Rad) and transferred to nitrocellulose membranes using the Trans-Blot turbo transfer system, (Bio-Rad). Primary antibodies (anti-RBP4, 1:2,000; #ab109193, Abcam, Cambridge, UK and anti-GAPDH, 1:40,000 #CB1001, Calbiochem, Merck-Millipore Amsterdam-Zuidoost, The Netherland) and horseradish peroxidase (HRP)-conjugated goat anti-rabbit secondary antibodies (1:2,000; Agilent DAKO, Amstelveen, The Netherlands) were used for detection. Proteins were detected using the Pierce ECL Western blotting kit (Thermo Fisher Scientific). Images were captured using the chemidoc XRS system and Image Lab version 3.0 (Bio-Rad). The intensity of bands was quantified using ImageJ version 1.51 (NIH, Maryland, USA).
Histology and pathological scoring
Hematoxylin and Eosin (H&E) staining’s on liver sections (4 µm) were performed. Snap-frozen liver sections (5 µm) were stained using Oil Red O (ORO) [20,21]. A pathological score was calculated as previously described [22] to determine the level of steatosis, as well as the grade of lobular inflammation.
CD68 staining
Immunohistochemistry for CD68 (1:300; rabbit anti-CD68; #137002, Biolegio, Nijmegen, the Netherlands) was performed on snap-frozen liver sections as previously described [23]. Antigen retrieval was performed by microwave irradiation in citrate buffer, pH 6.0 and blocking of endogenous peroxidase with 0.3% H2O2 for
30 min. Horseradish peroxidase (HRP)-conjugated goat anti-rabbit antibodies (1:50; #170-6515, Bio-Rad) was used secondary antibody. Slides were stained with 3,3'-Diaminobenzidine (DAB) for 10 min and Haematoxylin was used as a counter nuclear stain (2 min at RT). Finally, slides were dehydrated and mounted with Eukitt®,
(Sigma-Aldrich). Slides were scanned using a nanozoomer 2.0 HT digital slide scanner (C9600-12, Hamamatsu Photonics, Hamamatsu, Japan) and analyzed with Aperio ImageScope (version 11.1, Leica Microsystems, Amsterdam, The
149
Netherlands). CD68-positive cells were counted to assess the expansion of macrophages in the liver.
Lipid analysis
Lipids were extracted from 15% (w/v) liver homogenate in PBS by using the Bligh & Dyer method [24]. A colorimetric assay was used to determine total cholesterol (11489232, Roche Molecular Biochemicals, Almere, the Netherlands) and free cholesterol (113609910930, DiaSys Diagnostic Systems Gmbh, Holzheim, Germany). Cholesterol standards (DiaSys Diagnostic Systems Gmbh) were used as a reference. Triglycerides were quantified using the Trig/GB kit (1187771, Roche Molecular Biochemicals) and Roche Precimat Glycerol standards (16658800) were used as a reference.
Serum and hepatic vitamin A analysis
Serum and tissue vitamin A content was analyzed by reverse phase HPLC as previously described [25]. Briefly, tissue (30-50 mg) was homogenized in PBS to create a 15% (w/v) tissue homogenate. Then, tissue homogenate (66.70 µL equal to 10 mg of tissue) or serum (50 µL) were added in antioxidant mix (2.75 mL, containing 1153.39 mmol/L pyrogallol, 66.01 mmol/L butylated hydroxytoluene, 311.08 mmol/L ethylenediaminetetraacetic acid and 2064.71 mmol/L ascorbic acid, dissolved in methanol (2.67):dH20 (1), pH 5.4) and vortexed thoroughly for 1 min.
Retinol and retinyl esters were extracted and deproteinized twice with n-hexane in the presence of retinol acetate (100 µL, concentration 4 µmol/L) as an external standard to assess the level of recovery after the extraction procedure. Standard curves created from a range of concentrations of retinol and retinyl palmitate were used to determine absolute tissue and serum concentrations of these compounds. Additionally, two negative controls (only containing internal standard) and two positive controls (low and high concentrations of retinol plus internal standard) were included in each series of extractions. Samples were evaporated under N2 and
diluted in 300 µL 100% ultrapure ethanol. Then, 50 µL was injected in HPLC (Waters 2795 Alliance HT Separations Module, Connecticut, USA) for phase separation on a C18 column (Waters Symmetry C18, dimension 150 x 3.0 mm, particle size 5 µm, Waters Corporation, Milford, MA, USA) and measurement (UV-VIS, dual wavelength, UV-4075 Jasco, Tokyo, Japan). Retinoids in samples were identified by the exact
150
retention time of known standards in ultraviolet absorption at 325 nm by HPLC. Finally, retinol and retinyl palmitate concentrations were calculated and normalized to final volume or tissue weight.
Vitamin A autofluorescence in liver tissue
Autofluorescence analysis was performed on unstained cryostatic liver sections using a Zeiss LSM 780 NLO two-photon CLSM (Carl Zeiss, Jena, Germany) as previously described [26,27]. Briefly, cryostat liver sections were illuminated with an excitation filter of 366 nm band-pass interference, and spectra were recorded in the range of 400-680 nm with spectrum acquisition from 0.2 to 3 seconds.
Statistical analysis
Data is presented in group Mean±SEM and statistical analysis was performed using the GraphPad Prism 6 software package (GraphPad Software, San Diego, CA, USA). Statistical significance was determined by Mann-Whitney test (two groups), One-way ANOVA or Kruskal-Wallis (more than two groups) followed by post-hoc Dunns (compare all pairs of columns). P-values with ≤0.05*, ≤0.01**, ≤0.001*** were considered significant.
5.3. RESULTS
HFC diet increases body weight and liver fat content.
In order to study the effect of NAFLD on vitamin A metabolism, we fed mice a high-fat and high-cholesterol (HFC)-containing diet for 12 or 20 weeks and first confirmed the development and progression of fatty liver disease. As described by us [28,29] and other [30,31], the HFC diet induced significant body and liver weight gain after 20 weeks (Figure 1A, B). Hepatic total and free cholesterol, as well as total triglycerides, were significantly increased after 12 weeks of HFC-feeding and remained enhanced after 20 weeks (Figure 1C-E). Similarly, plasma total and free cholesterol, as well as insulin levels were significantly elevated HFC-fed mice (Figure
1F-H). H&E and Oil-Red-O (ORO) staining confirmed excessive fat accumulation in
the livers of HFC-fed mice and was associated with accumulation of CD68-positive inflammatory cells (Figure 1I).
151
Figure 1. Hepatic fat accumulation and steatohepatitis in HFC-fed mice.
Mice fed chow or HFC-diet for 12 or 20 weeks were analyzed for A) body weight, B) liver weight, C) liver total cholesterol levels, D) liver triglyceride levels, E) liver free cholesterol levels, F) plasma cholesterol levels, G) plasma free cholesterol levels, H) plasma insulin levels, I) H&E staining, Oil red O staining (ORO) and CD68 immunohistochemistry. Quantification of (immuno)histochemistry is described in Supplementary Methods and Materials.
152
HFC diet induces markers of hepatic lipid uptake and synthesis, inflammation and fibrosis in mice
Hepatic expression of genes involved in lipid uptake (Srb1, Cd36) and synthesis (Scd1, Fasn, Acc1, Srebp1c) were strongly induced by HFC-fed mice (Figure 2A). Moreover, the HFC diet lead to an early progression to steatohepatitis, with enhanced expression of inflammatory markers, including Cd68, Tnf-α, Nos2, Ccl2, Il-1β and Il-6 (Figure 2B) as well as the progressive induction of markers of fibrosis (Coll1a1, Acta2, Tgf-β, and Timp1) (Figure 2C).
153
Figure 2. Expression of hepatic genes involved in lipid uptake, lipid synthesis, inflammation and fibrosis in HFC-fed mice.
Mice fed chow or HFC-diet for 12 or 20 weeks were analyzed by Q-PCR for hepatic expression of genes involved in A) hepatic lipid uptake (Srb1, Cd36) and synthesis (Scd1, Fasn, Acc1), B) hepatic inflammation (Cd68, Tnf-α, Nos2, Ccl2, Il-1β, Il-6) and C) liver fibrosis (Coll1a1, Acta2, Tgf-β and Timp1). Hepatic lipid uptake and synthesis, hepatic inflammation and hepatic fibrosis
154
Reduced retinol, but elevated retinyl palmitate in livers of HFC-fed mice
Next, we determined the effect of the HFC diet on the levels of retinol in liver and plasma, as well as hepatic retinyl palmitate, the major storage form of vitamin A. Hepatic retinol levels were comparable in 12- and 20-week chow-fed mice (24.0 ± 2.4 and 24.0 ± 2.5 µg/g liver, respectively; Figure 3A). In contrast, hepatic retinol levels were progressively decreased in HFC-fed mice (9.1 ± 0.8 and 5.7 ± 0.8 µg/g liver after 12 and 20 weeks, respectively; Figure 3A). On the other hand, hepatic retinyl palmitate levels progressively increased in HFC-fed mice (785 ± 34 and 1,060 ± 182 µg/g liver after 12 and 20 weeks, respectively) and were significantly higher than in chow-fed mice (331 ± 50 and 413 ± 58 µg/g liver after 12 and 20 weeks, respectively;
Figure 3B). The disturbed hepatic retinol/retinyl palmitate balance was not
accompanied by changes in plasma levels of retinol after 12 weeks HFC diet (1.4 ± 0.1 and 1.6 ± 0.1 µmol/L for chow and HFC diet, respectively), while it was increased in HFC-fed mice after 20 weeks (1.1 ± 0.1 and 2.1 ± 0.1 µmol/L for chow and HFC diet, respectively; Figure 3C). Hepatic RBP4 protein levels were decreased in HFC-fed mice compared to chow-HFC-fed mice (Figure 3D), while hepatic mRNA expression of Rbp4 was comparable in all animal groups (Figure 4A). In contrast, serum RBP4 levels were elevated in HFC-fed mice, which was particularly evident after 20 weeks
(Figure 3E). These results show that a drastic reduction in hepatic retinol levels in
155
Figure 3. A HFC diet leads to impaired hepatic vitamin A metabolism in mice.
Mice fed chow or HFC-diet for 12 or 20 weeks were analyzed for hepatic levels of A) retinol B) retinyl palmitate, and C) plasma levels of retinol. Hepatic retinol levels were strongly reduced in HFC-fed mice, while retinyl palmitate levels were significantly increased compared to control mice. 12-week HFC-feeding did not alter plasma retinol levels, which were slightly elevated after 20 weeks. D) Hepatic RBP4 protein levels were decreased in 12 week HFC-fed mice, while this decrease was no longer significant after 20 week HFC feeding. E) Plasma RBP4 progressively increased in HFC- fed mice.
156
Hepatic expression of genes involved in vitamin A storage, transport, hydrolysis, as well as retinoic acid targets are elevated in HFC-fed mice
Hepatic mRNA levels of Lrat, coding for the main enzyme producing retinyl esters in the liver, increased with time in HFC-fed mice (Figure 4A). No or minor changes were detected in mRNA levels for alternative retinol-esterifying enzymes, like Dgat1 and Dgat2. Of the various retinyl ester hydrolases (REHs) that mobilize retinol from hepatic retinyl ester stores, only Adpn/Pnpla3 mRNA levels were strongly induced in HFC-fed mice, while Atgl/Pnpla2 and Hsl/Lipe mRNA levels were not altered compared to chow-fed mice. Raldh2 mRNA levels were significantly enhanced in livers of HFC-fed mice, while levels of other members of the retinol-to-retinoic-acid-converting enzymes [32–34] were not changed (Raldh1, Raldh3 and Raldh4) (Figure
4B). Hepatic expression of retinoic acid-responsive genes (RAR-β, Cyp26a1,
Hsd17b13, Ucp2, Cpt1a, Fgf21) was enhanced in the mice fed the HFC diet (Figure
4C). Thus, the HFC diet leads to a hyperdynamic state of retinol metabolism in mice,
157
Figure 4. A HFC diet strongly affects the hepatic expression of genes involved in vitamin A homeostasis.
Mice fed chow or HFC-diet for 12 or 20 weeks were analyzed by Q-PCR for hepatic expression of genes and transcription factors involved in A) vitamin A storage (Lrat, Dgat1, Dgat2) and transport (Rbp4), B) retinyl ester hydrolysis (Atgl/Pnpla2, Pnpla3, Lipe) and retinol-to-retinoic acid conversion (Raldh1, Raldh2, Raldh3, Raldh4) and C) retinoic acid target genes (Rar-β, Cyp26a1, Hsd17b13, Ucp2, Cpt1a, Fgf21). Hepatic expression of vitamin A storage and
hydrolysis were increased in HFC-fed mice, in conjunction with enhanced expression of retinoic acid responsive genes.
158
Reduced retinol and enhanced retinyl palmitate in livers of ob/ob mice
To validate our findings, we next analyzed hepatic vitamin A metabolism in another model of fatty liver disease, e.g. the leptin-deficient ob/ob mouse. In line with the observations described above for HFC-fed mice, hepatic retinol levels were significantly lower in ob/ob mice compared to age-matched wild-type littermates (2.8 ± 0.1 versus 16.9 ± 5.1 µg/g liver, respectively; Figure 5A), in conjunction with strongly enhanced levels of retinyl palmitate (359 ± 28 versus 160 ± 26 µg/g liver, respectively; Figure 5B). Moreover, plasma retinol and RBP4 levels were significantly higher in ob/ob mice as compared to wild-type littermates (2.5 ± 0.2 versus 0.9 ± 0.2 µM retinol, respectively; Figure 5C and D). Finally, also hepatic RBP4 protein levels were reduced in ob/ob mice independently of (unchanged) Rbp4 mRNA levels as compared to wild-type mice (Figure 5D and 6A), all features being comparable between HFC-fed wild type mice and chow-fed ob/ob mice.
Figure 5. Ob/ob mice show disturbed hepatic vitamin A metabolism.
Ob/ob mice and age-matched wild-type littermates were sacrificed and analyzed for A) hepatic
retinol, B) hepatic retinyl palmitate, C) plasma retinol and hepatic (D) and plasma (E) RBP4 levels. Hepatic retinol levels were strongly reduced, while retinyl palmitate levels were significantly increased in ob/ob mice. Plasma retinol and RBP4 protein levels were significantly elevated in ob/ob mice. In contrast, hepatic RBP4 protein levels were reduced. Note that hepatic
159
Hepatic expression of genes involved in vitamin A storage, transport, hydrolysis and retinoic acid-responsive targets are elevated in ob/ob mice
Similar to HFC-fed mice, hepatic expression of Lrat and Dgat1 were significantly enhanced in ob/ob mice as compared to controls, while expression of Dgat2 and Rbp4 was not different between both animal groups (Figure 6A). Pnpla3 was strongly enhanced in ob/ob mice compared to control mice, while Pnpla2 and Lipe levels were unchanged. In contrast to HFC-fed mice, Raldh2 mRNA levels were reduced in ob/ob mice, while Raldh1 and Raldh3 levels were significantly elevated
(Figure 6B). Finally, hepatic expression of various retinoic acid-responsive genes
was also enhanced in ob/ob mice livers as compared to controls (Figure 6C). Taken together, both NAFLD mouse models present comparable impairments in hepatic vitamin A homeostasis.
160
Figure 6. Hepatic expression of genes involved in vitamin A homeostasis is strongly affected in ob/ob mice.
Ob/ob mice and age-matched wild-type littermates were sacrificed and analyzed by Q-PCR for
hepatic expression of genes and transcription factors involved in A) vitamin A storage (Lrat,
Dgat1, Dgat2), transport (Rbp4), B) vitamin A hydrolysis (Atgl/Pnpla2, Pnpla3, Lipe) and
retinol-to-retinoic acid conversion (Raldh1, Raldh2, Raldh3, Raldh4) and C) retinoic acid target genes
(Rar-β, Cyp26a1, Hsd17b13, Ucp2, Cpt1a, Fgf21). Hepatic expression of vitamin A storage and
hydrolysis was increased in ob/ob, in conjunction with enhanced expression of retinoic acid responsive genes.
161
Palmitic acid increases Lrat expression and vitamin A accumulation in primary rat hepatocytes
Dietary vitamin A is absorbed by hepatocytes through retinyl ester-carrying chylomicron remnants, after which it is redistributed to HSC for storage (see introduction). Thus, both hepatocytes and HSC are possible sites for retinyl ester accumulation and earlier work has suggested vitamin A accumulation in hepatocytes of human and rodent fatty livers [26,35]. Microscopical analysis of liver tissue for vitamin A-specific autofluorescence revealed few bright (auto)fluorescent dots in the hepatic parenchyma of chow-fed mice (Figure 7A, top right panel) reminiscent of the distribution of quiescent HSC and the pattern observed by others in healthy rat and human liver [26,35]. The autofluorescence signal strongly increased in livers of HFC-fed mice, showing a completely altered staining pattern, which now appeared in large vesicular structures (“lipid droplets”) that localize predominantly in hepatocytes
(Figure 7A, bottom right panel).
In order to study cell-type specific effects of vitamin A storage in an in vitro model of fatty liver disease, we exposed primary rat hepatocytes, as well as quiescent and activated primary rat HSC, to palmitate and found that it enhanced Lrat mRNA levels only in hepatocytes and not in quiescent nor in activated HSC (Figure 7B). Conversely, PNPLA3 levels were elevated by palmitate exposure specifically in HSC. Moreover, cellular retinyl palmitate levels significantly increased only in primary rat hepatocytes exposed to palmitate together with retinol for 48 h (Figure 7C). These results suggest that NAFLD promotes vitamin A loss in HSC while vitamin A storage is induced in hepatocytes.
162
Figure 7. Autofluorescence in liver increases with fatty liver and palmitic acid induces
Lrat expression and cellular retinyl palmitate accumulation in primary rat hepatocytes.
Oil-Red-O staining (left panels) and vitamin A-specific autofluorescence (right panels of liver sections of chow-fed (top panels) and HFC-fed (bottom panels) mice. Vitamin A-specific autofluorescence was strongly increased in livers of HFC-fed mice and was located predominantly in hepatocytes compared to its location in sparsely-present HSC in livers of chow-fed mice. B) Freshly-isolated and 4 h-attached primary rat hepatocytes and quiescent primary HSC were and immediately treated with and without palmitate for 24 h. Cells were harvested and analyzed by Q-PCR analysis for Lrat and Pnpla3 expression. The gene expression is presented in 2-∆∆CT and normalized to 18S. C) Freshly-isolated and 4 h-attached primary rat hepatocytes were treated for 48 h with and without palmitate and retinol, followed by an analysis of cellular retinyl palmitate and retinol levels.
163
5.4. DISCUSSION
In this study, we show that steatohepatitis in mice heavily affects hepatic vitamin A metabolism leading to reduced retinol and enhanced retinyl ester levels in the liver. Notably, retinyl esters accumulate in hepatocytes rather than in hepatic stellate cells, cells that store vitamin A esters in a healthy liver. Following retinol levels, hepatic RBP4 levels are also reduced, while increased in circulation. Thus, steatohepatitis does not lead to true vitamin A deficiency as suggested by earlier studies, but rather leads to hepatic retinol deficiency due to metabolic changes. Vitamin A supplementation in the form of retinyl esters may, therefore, be counterproductive in NAFLD, as it will likely accumulate in the already overloaded hepatocytes. Also, aberrant vitamin A metabolism may directly affect the activity of nuclear receptors, such as RAR, PPARs, LXR and FXR, as they require RXR for most of their actions. Chronic liver diseases, including NAFLD, are associated with low serum and hepatic retinol levels, which is generally considered to be a sign of systemic vitamin A deficiency [10,11,36–38]. Moreover, serum and hepatic retinol levels are negatively correlated with liver disease progression [10,36,39,40]. However, systemic retinol levels are only a small fraction of the total pool of vitamin A, which mostly consists of retinyl esters stored in liver (≥80%) and adipose tissue (10-20%) [41,42]. Thus, systemic retinol levels alone are not a reliable marker for hypovitaminosis A per se. Earlier studies have actually suggested that NAFLD is associated with hepatic vitamin A accumulation in humans and rodents [11,43].
We show that hepatic fat accumulation in mice is associated with a strong reduction in hepatic retinol levels, but not in circulation. This is in line with earlier observations [10,11,36–38] and indicates that the mouse NAFLD models used do show the hepatic phenotype, but do not replicate the reduced circulatory retinol levels observed in NAFLD patients [10,36,44]. NAFLD mouse models likely reflect only the initiating phase of NAFLD and serum retinol levels are also hardly affected at that stage in patients. Alternatively, the observed difference may be species related, but this requires analyses of vitamin A metabolism in more severe mouse NAFLD models. We did observe, however, that serum RBP4 levels are elevated in NAFLD mice, which is also found in obese individuals with or without established NAFLD [45–49]. In contrast, hepatic RBP4 levels are reduced in NAFLD mice. This implies that, even though hepatic retinol levels are low, sufficient retinol is produced to promote -and even enhance- RBP4 secretion from the liver. Efficient RBP4 secretion
164
from the liver strongly depends on retinol availability. Vitamin A deficiency leads to pronounced hepatic accumulation of retinol-free apo-RBP4 under unchanged Rbp4 mRNA levels and is rapidly released into the circulation upon retinol or retinoic acid treatment [50–52]. As hepatic retinol levels were low, enhanced RBP4 secretion may also result from enhanced production of retinoic acids. Indeed, mRNA levels of Raldh1, the main enzyme responsible for RA production in the liver, were significantly elevated in ob/ob mice, with similar trends observed for 12- and 20-wk HFC-fed mice. Moreover, expression of various RA-responsive genes, such as RAR-β, Cyp26a1, Hsd17b13, Ucp2, Cpt1a, Fgf21, was also induced in NAFLD mice. This is in line with the earlier reported hyper-metabolic state of hepatic vitamin A metabolism and degradation in NAFLD patients [13]. That study also observed enhanced expression of hepatic LRAT and DGAT1 in NAFLD patients, genes that encode enzymes that esterify retinol, which we also detected in NAFLD mice. This coincides with a strong increase in hepatic retinyl palmitate levels in NAFLD mice, which is the main retinyl ester in both human and rodents [53,54]. Particularly relevant is the apparent redistribution of vitamin A from HSC in control livers to lipid-loaded hepatocytes in fatty livers. Retinyl esters are rapidly converted to retinol in the healthy liver. Accumulation of retinyl esters in lipid-loaded hepatocytes may, therefore, result from decreased hydrolysis and/or increased esterification. Expression of Atgl/Pnpla2 (hydrolysis) was not changed, while both Lrat and Dgat1 (esterification) were enhanced in mouse fatty livers, indicating a shift to vitamin A storage in retinyl esters. On the other hand, PNPLA3/ADPN is also able to hydrolyze retinyl esters [55] and its expression is strongly increased in mouse fatty liver, like in human NAFLD patients [56]. Still, PNPLA3’s main substrates are TG that also strongly accumulate in mouse fatty liver. This implies that its activity cannot compensate for the build-up of TG in hepatocytes, as observed for retinyl palmitate. Notably, Lrat expression was induced in primary rat hepatocytes exposed to palmitate and they accumulate retinyl-palmitate when co-exposed to retinol. These conditions did not regulate Lrat expression in qHSC, but rather enhance retinyl ester hydrolysis by increasing PNPLA3 expression. These findings are in line with the shift in the main location of vitamin A to hepatocytes in fatty liver in mice (this study), rats [35] and human [26]. The redistribution of retinyl esters from HSC to hepatocytes may also promote NAFLD-associated fibrosis, as differentiation of qHSC to myofibroblasts is characterized by the loss of vitamin A-containing lipid droplets. In NAFLD, this
165
apparently happens in conjunction with a pronounced accumulation of retinyl esters in the liver, but then primarily in hepatocytes.
Our observation of retinyl palmitate accumulation in mouse fatty liver contrasts to recent findings by others, who reported pronounced reductions in hepatic retinol and retinyl palmitate in HFD-fed mice and ob/ob mice [11]. Several reasons may account for this discrepancy; 1) HFD vs HFC-diets: A high-fat (only) diet may affect vitamin A metabolism differently than the high-fat high-cholesterol diets (HFC) used in this study. However, we did not detect a reduction in hepatic retinyl palmitate levels in mice fed HFD diet for 12 weeks, while hepatic retinol levels were reduced in both HFD- and HFC-fed mice (Supplementary Figure S1 and Figure 3A). 2) Amount of vitamin A in the diets: In our study, both control chow and HFC diet contained equal amounts of vitamin A, e.g. 20 IU/g. The earlier study [11] used a HFD that contained significantly less vitamin A compared to the control chow (3.8 vs. 15 IU/g plus additional β-carotene, respectively). Dietary intake of vitamin A, even above daily recommendations, correlates directly with hepatic levels of retinyl palmitate and retinol [57–60]. Thus, for establishing an effect of a (high-fat) diet on hepatic vitamin A metabolism it is crucial to standardize the dietary vitamin A intake for control and experimental diet. This is the case in the ob/ob mouse studies and cannot explain the opposing results between our and the earlier study. 3) Retinol/retinyl ester extraction procedure: extraction of retinol and retinyl esters from serum or tissue is typically performed with either n-hexane [25,41] or acetonitrile [11], though this is not always specified in methods sections. We compared both methods and found that acetonitrile very inefficiently extracts retinyl palmitate specifically from fatty liver tissue, while retinol extraction was similar with these solvents (Supplementary
Figure S2). This suggests that the extraction of retinyl esters by acetonitrile is
compromised when a lot of fat is present in the (liver) tissue, and this may (in part) explain the controversial findings on retinyl ester levels in fatty liver.
Thus, our study shows that fatty liver disease is associated with hepatic accumulation of retinyl esters and disturbed vitamin A metabolism. The latter condition likely modulates disease progression, as it was recently shown that even moderate changes in hepatic RA production significantly enhance hepatic lipid accumulation [61]. Moreover, lipid metabolism is tightly controlled by various nuclear receptors (NR), like PPARs, FXR, LXR and RAR, ligands of which are in advanced clinical trial stages for the treatment of NAFLD [62]. All these factors require RXR as an obligate partner and aberrant production of RXR-activating retinoids will affect NR/RXR
166
signaling. Further studies are needed to determine the impact of changed vitamin A metabolism in NAFLD on the therapeutic efficacy of these drug targets.
ACKNOWLEDGEMENTS
The authors want to acknowledge J.W. (Hans) Jonker (group), Tim Van Zutphen and Dicky Struik for providing ob/ob mice materials.
167
REFERENCES
[1] S. Albhaisi, A. Sanyal, Recent advances in understanding and managing non-alcoholic fatty liver disease, F1000Research. 7 (2018). doi:10.12688/f1000research.14421.1.
[2] S. Bellentani, F. Scaglioni, M. Marino, G. Bedogni, Epidemiology of Non-Alcoholic Fatty Liver Disease, Dig. Dis. 28 (2010) 155–161. doi:10.1159/000282080.
[3] C.N. Katsagoni, M. Georgoulis, G.V. Papatheodoridis, D.B. Panagiotakos, M.D. Kontogianni, Effects of lifestyle interventions on clinical characteristics of patients with non-alcoholic fatty liver disease: A meta-analysis, Metabolism. 68 (2017) 119–132. doi:10.1016/j.metabol.2016.12.006.
[4] J.P. Iredale, A. Thompson, N.C. Henderson, Extracellular matrix degradation in liver fibrosis: Biochemistry and regulation, Biochim. Biophys. Acta. 1832 (2013) 876–883. doi:10.1016/j.bbadis.2012.11.002.
[5] R. Schreiber, U. Taschler, K. Preiss-Landl, N. Wongsiriroj, R. Zimmermann, A. Lass, Retinyl ester hydrolases and their roles in vitamin A homeostasis, Biochim. Biophys. Acta. 1821 (2012) 113–123. doi:10.1016/j.bbalip.2011.05.001.
[6] W.S. Blaner, S.M. O’Byrne, N. Wongsiriroj, J. Kluwe, D.M. D’Ambrosio, H. Jiang, R.F. Schwabe, E.M.C. Hillman, R. Piantedosi, J. Libien, Hepatic stellate cell lipid droplets: a specialized lipid droplet for retinoid storage, Biochim. Biophys. Acta. 1791 (2009) 467–473. doi:10.1016/j.bbalip.2008.11.001. [7] H. Senoo, Y. Mezaki, M. Fujiwara, The stellate cell system (vitamin A-storing
cell system), Anat. Sci. Int. 92 (2017) 387–455. doi:10.1007/s12565-017-0395-9.
[8] A. Saeed, M. Hoekstra, M.O. Hoeke, J. Heegsma, K.N. Faber, The interrelationship between bile acid and vitamin A homeostasis, Biochim. Biophys. Acta. 1862 (2017) 496–512. doi:10.1016/j.bbalip.2017.01.007. [9] World Health Organization, Indicators for assessing vitamin A deficiency and
their application in monitoring and evaluating intervention programmes, Eileen Brown, James Akré, World Health Organization: Geneva, Switzerland, 1996. http://www.who.int/nutrition/publications/micronutrients/vitamin_a_deficiency/W HO_NUT_96.10/en/.
[10] G.V. Chaves, S.E. Pereira, C.J. Saboya, D. Spitz, C.S. Rodrigues, A. Ramalho, Association between liver vitamin A reserves and severity of nonalcoholic fatty liver disease in the class III obese following bariatric surgery, Obes. Surg. 24 (2014) 219–224. doi:10.1007/s11695-013-1087-8.
[11] S.E. Trasino, X.-H. Tang, J. Jessurun, L.J. Gudas, Obesity Leads to Tissue, but not Serum Vitamin A Deficiency, Sci. Rep. 5 (2015) 15893. doi:10.1038/srep15893.
[12] M. Kovarova, I. Königsrainer, A. Königsrainer, F. Machicao, H.-U. Häring, E. Schleicher, A. Peter, The Genetic Variant I148M in PNPLA3 Is Associated With Increased Hepatic Retinyl-Palmitate Storage in Humans, J. Clin. Endocrinol. Metab. 100 (2015) E1568-1574. doi:10.1210/jc.2015-2978. [13] A.A. Ashla, Y. Hoshikawa, H. Tsuchiya, K. Hashiguchi, M. Enjoji, M.
Nakamuta, A. Taketomi, Y. Maehara, K. Shomori, A. Kurimasa, I. Hisatome, H. Ito, G. Shiota, Genetic analysis of expression profile involved in retinoid metabolism in non-alcoholic fatty liver disease, Hepatol. Res. Off. J. Jpn. Soc. Hepatol. 40 (2010) 594–604. doi:10.1111/j.1872-034X.2010.00646.x.
168
[14] T.E. Vrenken, M. Buist-Homan, A.J. Kalsbeek, K.N. Faber, H. Moshage, The active metabolite of leflunomide, A77 1726, protects rat hepatocytes against bile acid-induced apoptosis, J. Hepatol. 49 (2008) 799–809. doi:10.1016/j.jhep.2008.07.019.
[15] H. Moshage, A. Casini, C.S. Lieber, Acetaldehyde selectively stimulates collagen production in cultured rat liver fat-storing cells but not in hepatocytes, Hepatol. Baltim. Md. 12 (1990) 511–518.
[16] S. Shajari, A. Laliena, J. Heegsma, M.J. Tuñón, H. Moshage, K.N. Faber, Melatonin suppresses activation of hepatic stellate cells through RORα-mediated inhibition of 5-lipoxygenase, J. Pineal Res. 59 (2015) 391–401. doi:10.1111/jpi.12271.
[17] Y.M. Song, S.-O. Song, Y.-K. Jung, E.-S. Kang, B.S. Cha, H.C. Lee, B.-W. Lee, Dimethyl sulfoxide reduces hepatocellular lipid accumulation through autophagy induction, Autophagy. 8 (2012) 1085–1097. doi:10.4161/auto.20260.
[18] H. Blokzijl, S. Vander Borght, L.I.H. Bok, L. Libbrecht, M. Geuken, F.A.J. van den Heuvel, G. Dijkstra, T.A.D. Roskams, H. Moshage, P.L.M. Jansen, K.N. Faber, Decreased P-glycoprotein (P-gp/MDR1) expression in inflamed human intestinal epithelium is independent of PXR protein levels, Inflamm. Bowel Dis. 13 (2007) 710–720. doi:10.1002/ibd.20088.
[19] A. Pellicoro, F.A.J. van den Heuvel, M. Geuken, H. Moshage, P.L.M. Jansen, K.N. Faber, Human and rat bile acid-CoA:amino acid N-acyltransferase are liver-specific peroxisomal enzymes: implications for intracellular bile salt transport, Hepatol. Baltim. Md. 45 (2007) 340–348. doi:10.1002/hep.21528. [20] A.H. Fischer, K.A. Jacobson, J. Rose, R. Zeller, Hematoxylin and eosin
staining of tissue and cell sections, CSH Protoc. 2008 (2008) pdb.prot4986. [21] A. Mehlem, C.E. Hagberg, L. Muhl, U. Eriksson, A. Falkevall, Imaging of
neutral lipids by oil red O for analyzing the metabolic status in health and disease, Nat. Protoc. 8 (2013) 1149–1154. doi:10.1038/nprot.2013.055. [22] D.E. Kleiner, E.M. Brunt, M. Van Natta, C. Behling, M.J. Contos, O.W.
Cummings, L.D. Ferrell, Y.-C. Liu, M.S. Torbenson, A. Unalp-Arida, M. Yeh, A.J. McCullough, A.J. Sanyal, Nonalcoholic Steatohepatitis Clinical Research Network, Design and validation of a histological scoring system for nonalcoholic fatty liver disease, Hepatol. Baltim. Md. 41 (2005) 1313–1321. doi:10.1002/hep.20701.
[23] M. Aparicio-Vergara, P.P.H. Hommelberg, M. Schreurs, N. Gruben, R. Stienstra, R. Shiri-Sverdlov, N.J. Kloosterhuis, A. de Bruin, B. van de Sluis, D.P.Y. Koonen, M.H. Hofker, Tumor necrosis factor receptor 1 gain-of-function mutation aggravates nonalcoholic fatty liver disease but does not cause insulin resistance in a murine model, Hepatol. Baltim. Md. 57 (2013) 566–576. doi:10.1002/hep.26046.
[24] E.G. Bligh, W.J. Dyer, A rapid method of total lipid extraction and purification, Can. J. Biochem. Physiol. 37 (1959) 911–917. doi:10.1139/o59-099.
[25] Y.-K. Kim, L. Quadro, Reverse-phase high-performance liquid chromatography (HPLC) analysis of retinol and retinyl esters in mouse serum and tissues, Methods Mol. Biol. Clifton NJ. 652 (2010) 263–275. doi:10.1007/978-1-60327-325-1_15.
[26] A.C. Croce, U. De Simone, I. Freitas, E. Boncompagni, D. Neri, U. Cillo, G. Bottiroli, Human liver autofluorescence: an intrinsic tissue parameter discriminating normal and diseased conditions, Lasers Surg. Med. 42 (2010) 371–378. doi:10.1002/lsm.20923.
169
[27] A.C. Croce, U. De Simone, M. Vairetti, A. Ferrigno, E. Boncompagni, I. Freitas, G. Bottiroli, Liver autofluorescence properties in animal model under altered nutritional conditions, Photochem. Photobiol. Sci. Off. J. Eur. Photochem. Assoc. Eur. Soc. Photobiol. 7 (2008) 1046–1053. doi:10.1039/b804836c. [28] P. Bartuzi, T. Wijshake, D.C. Dekker, A. Fedoseienko, N.J. Kloosterhuis, S.A.
Youssef, H. Li, R. Shiri-Sverdlov, J.-A. Kuivenhoven, A. de Bruin, E. Burstein, M.H. Hofker, B. van de Sluis, A cell-type-specific role for murine Commd1 in liver inflammation, Biochim. Biophys. Acta. 1842 (2014) 2257–2265. doi:10.1016/j.bbadis.2014.06.035.
[29] P. Bartuzi, D.D. Billadeau, R. Favier, S. Rong, D. Dekker, A. Fedoseienko, H. Fieten, M. Wijers, J.H. Levels, N. Huijkman, N. Kloosterhuis, H. van der Molen, G. Brufau, A.K. Groen, A.M. Elliott, J.A. Kuivenhoven, B. Plecko, G. Grangl, J. McGaughran, J.D. Horton, E. Burstein, M.H. Hofker, B. van de Sluis, CCC- and WASH-mediated endosomal sorting of LDLR is required for normal clearance of circulating LDL, Nat. Commun. 7 (2016) 10961. doi:10.1038/ncomms10961. [30] R. Costa, I. Rodrigues, L. Guardão, S. Rocha-Rodrigues, C. Silva, J.
Magalhães, M. Ferreira-de-Almeida, R. Negrão, R. Soares, Xanthohumol and 8-prenylnaringenin ameliorate diabetic-related metabolic dysfunctions in mice, J. Nutr. Biochem. 45 (2017) 39–47. doi:10.1016/j.jnutbio.2017.03.006.
[31] M.S. Kim, M.-S. Choi, S.N. Han, High fat diet-induced obesity leads to proinflammatory response associated with higher expression of NOD2 protein, Nutr. Res. Pract. 5 (2011) 219–223. doi:10.4162/nrp.2011.5.3.219.
[32] N.L. Makia, P. Bojang, K.C. Falkner, D.J. Conklin, R.A. Prough, Murine hepatic aldehyde dehydrogenase 1a1 is a major contributor to oxidation of aldehydes formed by lipid peroxidation, Chem. Biol. Interact. 191 (2011) 278–287. doi:10.1016/j.cbi.2011.01.013.
[33] E.C. Kathmann, S. Naylor, J.J. Lipsky, Rat liver constitutive and phenobarbital-inducible cytosolic aldehyde dehydrogenases are highly homologous proteins that function as distinct isozymes, Biochemistry. 39 (2000) 11170–11176. [34] D.K. Bhatt, A. Gaedigk, R.E. Pearce, J.S. Leeder, B. Prasad, Age-dependent
Protein Abundance of Cytosolic Alcohol and Aldehyde Dehydrogenases in Human Liver, Drug Metab. Dispos. Biol. Fate Chem. 45 (2017) 1044–1048. doi:10.1124/dmd.117.076463.
[35] A.C. Croce, A. Ferrigno, V.M. Piccolini, E. Tarantola, E. Boncompagni, V. Bertone, G. Milanesi, I. Freitas, M. Vairetti, G. Bottiroli, Integrated autofluorescence characterization of a modified-diet liver model with accumulation of lipids and oxidative stress, BioMed Res. Int. 2014 (2014) 803491. doi:10.1155/2014/803491.
[36] J.I. Botella-Carretero, J.A. Balsa, C. Vázquez, R. Peromingo, M. Díaz-Enriquez, H.F. Escobar-Morreale, Retinol and alpha-tocopherol in morbid obesity and nonalcoholic fatty liver disease, Obes. Surg. 20 (2010) 69–76. doi:10.1007/s11695-008-9686-5.
[37] P.V. Havaldar, V.D. Patel, B.M. Siddibhavi, Recurrent vitamin A deficiency and fatty liver, J. Trop. Pediatr. 37 (1991) 87–88.
[38] Y. Liu, H. Chen, J. Wang, W. Zhou, R. Sun, M. Xia, Association of serum retinoic acid with hepatic steatosis and liver injury in nonalcoholic fatty liver disease, Am. J. Clin. Nutr. 102 (2015) 130–137. doi:10.3945/ajcn.114.105155. [39] L. de Souza Valente da Silva, G. Valeria da Veiga, R.A. Ramalho, Association
of serum concentrations of retinol and carotenoids with overweight in children and adolescents, Nutr. Burbank Los Angel. Cty. Calif. 23 (2007) 392–397. doi:10.1016/j.nut.2007.02.009.
170
[40] M.L. Neuhouser, C.L. Rock, A.L. Eldridge, A.R. Kristal, R.E. Patterson, D.A. Cooper, D. Neumark-Sztainer, L.J. Cheskin, M.D. Thornquist, Serum concentrations of retinol, alpha-tocopherol and the carotenoids are influenced by diet, race and obesity in a sample of healthy adolescents, J. Nutr. 131 (2001) 2184–2191.
[41] M.A. Kane, A.E. Folias, J.L. Napoli, HPLC/UV quantitation of retinal, retinol, and retinyl esters in serum and tissues, Anal. Biochem. 378 (2008) 71–79. doi:10.1016/j.ab.2008.03.038.
[42] W.S. Blaner, Y. Li, P.-J. Brun, J.J. Yuen, S.-A. Lee, R.D. Clugston, Vitamin A Absorption, Storage and Mobilization, Subcell. Biochem. 81 (2016) 95–125. doi:10.1007/978-94-024-0945-1_4.
[43] S.E. Trasino, X.-H. Tang, J. Jessurun, L.J. Gudas, Retinoic acid receptor β2 agonists restore glycaemic control in diabetes and reduce steatosis, Diabetes Obes. Metab. 18 (2016) 142–151. doi:10.1111/dom.12590.
[44] F.I. Suano de Souza, O.M. Silverio Amancio, R.O. Saccardo Sarni, T. Sacchi Pitta, A.P. Fernandes, F.L. Affonso Fonseca, S. Hix, R.A. Ramalho, Non-alcoholic fatty liver disease in overweight children and its relationship with retinol serum levels, Int. J. Vitam. Nutr. Res. Int. Z. Vitam.- Ernahrungsforschung J. Int. Vitaminol. Nutr. 78 (2008) 27–32. doi:10.1024/0300-9831.78.1.27.
[45] Q. Yang, T.E. Graham, N. Mody, F. Preitner, O.D. Peroni, J.M. Zabolotny, K. Kotani, L. Quadro, B.B. Kahn, Serum retinol binding protein 4 contributes to insulin resistance in obesity and type 2 diabetes, Nature. 436 (2005) 356–362. doi:10.1038/nature03711.
[46] D.G. Haider, K. Schindler, G. Prager, A. Bohdjalian, A. Luger, M. Wolzt, B. Ludvik, Serum retinol-binding protein 4 is reduced after weight loss in morbidly obese subjects, J. Clin. Endocrinol. Metab. 92 (2007) 1168–1171. doi:10.1210/jc.2006-1839.
[47] T. Reinehr, B. Stoffel-Wagner, C.L. Roth, Retinol-binding protein 4 and its relation to insulin resistance in obese children before and after weight loss, J. Clin. Endocrinol. Metab. 93 (2008) 2287–2293. doi:10.1210/jc.2007-2745. [48] M. Tajtáková, Z. Semanová, G. Ivancová, J. Petrovicová, V. Donicová, E.
Zemberová, [Serum level of retinol-binding protein 4 in obese patients with insulin resistance and in patients with type 2 diabetes treated with metformin], Vnitr. Lek. 53 (2007) 960–963.
[49] F. Ulgen, C. Herder, M.C. Kühn, H.S. Willenberg, M. Schott, W.A. Scherbaum, S. Schinner, Association of serum levels of retinol-binding protein 4 with male sex but not with insulin resistance in obese patients, Arch. Physiol. Biochem. 116 (2010) 57–62. doi:10.3109/13813451003631421.
[50] H. Ronne, C. Ocklind, K. Wiman, L. Rask, B. Obrink, P.A. Peterson, Ligand-dependent regulation of intracellular protein transport: effect of vitamin a on the secretion of the retinol-binding protein, J. Cell Biol. 96 (1983) 907–910. [51] J.L. Dixon, D.S. Goodman, Studies on the metabolism of retinol-binding protein
by primary hepatocytes from retinol-deficient rats, J. Cell. Physiol. 130 (1987) 14–20. doi:10.1002/jcp.1041300104.
[52] D. Bellovino, Y. Lanyau, I. Garaguso, L. Amicone, C. Cavallari, M. Tripodi, S. Gaetani, MMH cells: An in vitro model for the study of retinol-binding protein secretion regulated by retinol, J. Cell. Physiol. 181 (1999) 24–32. doi:10.1002/(SICI)1097-4652(199910)181:1<24::AID-JCP3>3.0.CO;2-0.
171
[53] S. Futterman, J.S. Andrews, The Composition of Liver Vitamin A Ester and the Synthesis of Vitamin A Ester by Liver Microsomes, J. Biol. Chem. 239 (1964) 4077–4080.
[54] M.W. Schäffer, S.S. Roy, S. Mukherjee, D. Nohr, M. Wolter, H.K. Biesalski, D.E. Ong, S.K. Das, Qualitative and quantitative analysis of retinol, retinyl esters, tocopherols and selected carotenoids out of various internal organs form different species by HPLC, Anal. Methods Adv. Methods Appl. 2 (2010) 1320–1332. doi:10.1039/c0ay00288g.
[55] C. Pirazzi, L. Valenti, B.M. Motta, P. Pingitore, K. Hedfalk, R.M. Mancina, M.A. Burza, C. Indiveri, Y. Ferro, T. Montalcini, C. Maglio, P. Dongiovanni, S. Fargion, R. Rametta, A. Pujia, L. Andersson, S. Ghosal, M. Levin, O. Wiklund, M. Iacovino, J. Borén, S. Romeo, PNPLA3 has retinyl-palmitate lipase activity in human hepatic stellate cells, Hum. Mol. Genet. 23 (2014) 4077–4085. doi:10.1093/hmg/ddu121.
[56] G. Aragonès, T. Auguet, S. Armengol, A. Berlanga, E. Guiu-Jurado, C. Aguilar, S. Martínez, F. Sabench, J.A. Porras, M.D. Ruiz, M. Hernández, J.J. Sirvent, D. Del Castillo, C. Richart, PNPLA3 Expression Is Related to Liver Steatosis in Morbidly Obese Women with Non-Alcoholic Fatty Liver Disease, Int. J. Mol. Sci. 17 (2016). doi:10.3390/ijms17050630.
[57] P.R. Sundaresan, S.M. Kaup, P.W. Wiesenfeld, S.J. Chirtel, S.C. Hight, J.I. Rader, Interactions in indices of vitamin A, zinc and copper status when these nutrients are fed to rats at adequate and increased levels, Br. J. Nutr. 75 (1996) 915–928.
[58] I. Johansson, G. Hallmans, A. Wikman, C. Biessy, E. Riboli, R. Kaaks, Validation and calibration of food-frequency questionnaire measurements in the Northern Sweden Health and Disease cohort, Public Health Nutr. 5 (2002) 487–496. doi:10.1079/PHNPHN2001315.
[59] C.J. Cifelli, A.C. Ross, Chronic vitamin A status and acute repletion with retinyl palmitate are determinants of the distribution and catabolism of all-trans-retinoic acid in rats, J. Nutr. 137 (2007) 63–70.
[60] A. Olivares, A. Daza, A.I. Rey, C.J. López-Bote, Dietary vitamin A concentration alters fatty acid composition in pigs, Meat Sci. 81 (2009) 295– 299. doi:10.1016/j.meatsci.2008.07.029.
[61] D. Yang, M.G. Vuckovic, C.P. Smullin, M. Kim, C. Pui-See Lo, E. Devericks, H.S. Yoo, M. Tintcheva, Y. Deng, J.L. Napoli, Modest Decreases in Endogenous All-Trans-Retinoic Acid Produced by a MouseRdh10Heterozygote Provoke Major Abnormalities in Adipogenesis and Lipid Metabolism, Diabetes. (2018). doi:10.2337/db17-0946.
[62] A. Saeed, R.P.F. Dullaart, T.C.M.A. Schreuder, H. Blokzijl, K.N. Faber, Disturbed Vitamin A Metabolism in Non-Alcoholic Fatty Liver Disease (NAFLD), Nutrients. 10 (2017). doi:10.3390/nu10010029.
172
Supplementary Figure 1: A high-fat diet (HFD) leads to impaired hepatic vitamin A metabolism in mice.
Mice were fed a chow diet or HFD diet for 12 weeks and analyzed for hepatic levels of retinol hepatic levels of retinyl palmitate. Hepatic retinol levels were strongly reduced in HFD-fed mice as compared to control mice. However, hepatic retinyl palmitate levels did change in both groups.
Supplementary Figure 2: Comparison n-hexane and ACN extraction in the mice liver.
Effect of two different methods of vitamin A extraction was analyzed with two different extraction methods 1) n-hexane (n-hex) (used in this study or 2) acetonitrile method (ACN) as previously described (Trasino et al. 2015. [11]) from same mice livers fed a chow diet or HFC diet for 20 weeks (n=3). Remarkably, both methods extracted a similar amount of retinol, but a significant less fraction of retinyl esters were extracted with ACN method.
173
Supplementary Table S1: Primers and probes used in this study.
Gene / ID Taqman primers and probe
36B4 NM_022402 Fwd: 5'-GCTTCATTGTGGGAGCAGACA-3' Rev: 5'-CATGGTGTTCTTGCCCATCAG-3' Probe: 5'-TCCAAGCAGATGCAGCAGATCCGC-3' Acaca / Acc1 NM_133360.1 / NM_022193.1 Fwd: 5'-GCCATTGGTATTGGGGCTTAC-3' Rev: 5'-CCCGACCAAGGACTTTGTTG-3' Probe: 5'-CTCAACCTGGATGGTTCTTTGTCCCAGC-3' Act2 / α-Sma NM_007392 Fwd: 5'-TTCGTGTGGCCCCTGAAG-3' Rev: 5'-GGACAGCACAGCCTGAATAGC-3' Probe: 5'-TTGAGACCTTCAATGTCCCCGCCA-3' Cd36 BC010262 / NM_031561 Fwd: 5'-GATCGGAACTGTGGGCTCAT-3' Rev: 5'-GGTTCCTTCTTCAAGGACAACTTC-3' Probe: 5'-AGAATGCCTCCAAACACAGCCAGGAC-3' Cd68 NM_009853 Fwd: 5'-CACTTCGGGCCATGTTTCTC-3' Rev: 5'-AGGACCAGGCCAATGATGAG-3' Probe: 5'-CAACCGTGACCAGTCCCTCTTGCTG-3' Ccl2 NM_031530.1 Fwd: 5'-TGTCTCAGCCAGATGCAGTTAAT-3' Rev: 5'-CCGACTCATTGGGATCATCTT-3' Probe: 5'-CCCCACTCACCTGCTGCTACTCATTCA-3' Col1a1 NM_007742 Fwd: 5'-TGGTGAACGTGGTGTACAAGGT-3' Rev: 5'-CAGTATCACCCTTGGCACCAT-3' Probe: 5'-TCCTGCTGGTCCCCGAGGAAACA-3' Cpt1a NM_013495.1 Fwd: 5'-CTCAGTGGGAGCGACTCTTCA-3' Rev: 5'-GGCCTCTGTGGTACACGACAA-3' Probe: 5'-CCTGGGGAGGAGACAGACACCATCCAAC-3' Mlxipl / Chrebp NM_021455.3 / NM_133552.1 Fwd: 5'-GATGGTGCGAACAGCTCTTCT-3' Rev: 5'-CTGGGCTGTGTCATGGTGAA-3' Probe: 5'-CCAGGCTCCTCCTCGGAGCCC-3' Cyp26a1 NM_007811.1 Fwd: 5'-GGAGACCCTGCGATTGAATC-3' Rev: 5'-GATCTGGTATCCATTCAGCTCAAA-3' Probe: 5'-TCTTCAGAGCAACCCGAAACCCTCC-3' Dgat1 NM_010046.2 / NM_053437.1 Fwd: 5'-GGTGCCCTGACAGAGCAGAT-3' Rev: 5'-CAGTAAGGCCACAGCTGCTG-3' Probe: 5'-CTGCTGCTACATGTGGTTAACCTGGCCA-3' Dgat2 NM_026384.2 / NM_001012345.1 Fwd: 5'-GGGTCCAGAAGAAGTTCCAGAAG-3' Rev: 5'-CCCAGGTGTCAGAGGAGAAGAG-3' Probe: 5'-CCCCTGCATCTTCCATGGCCG-3' Fasn
NM_007988 / NM_017332 Fwd: 5'-GGCATCATTGGGCACTCCTT-3' Rev: 5'-GCTGCAAGCACAGCCTCTCT-3'
Probe: 5'-CCATCTGCATAGCCACAGGCAACCTC-3' Fgf21 NM_020013.4 / NM_130752.1 Fwd: 5'-CCGCAGTCCAGAAAGTCTCC-3' Rev: 5'-TGACACCCAGGATTTGAATGAC-3' Probe: 5'-CCTGGCTTCAAGGCTTTGAGCTCC A-3'
Hsd17b13 NM_198030.2 / NM_001163486.1 Fwd: 5'- AAAGCAGAAAAGCAGACTGGTTCT-3' Rev: 5'- CCCCAGTTTCCTGCATTTGT-3' Probe: 5'-CGGTTTCCTCAACACCACGCTTATTGA-3' Lipe / HSL NM_010719 / X51415 Fwd: 5'-GAGGCCTTTGAGATGCCACT-3' Rev: 5'-AGATGAGCCTGGCTAGCACAG-3' Probe: 5'-CCATCTCACCTCCCTTGGCACACAC-3' IL-1β NM_008361 Fwd: 5'-ACCCTGCAGCTGGAGAGTGT-3' Rev: 5'-TTGACTTCTATCTTGTTGAAGACAAACC-3' Probe: 5'-CCCAAGCAATACCCAAAGAAGAAGATGGAA -3' IL-6 NM_031168 Fwd: 5'-CCGGAGAGGAGACTTCACAGA-3' Rev: 5'-AGAATTGCCATTGCACAACTCTT-3'
174
Probe: 5'-ACCACTTCACAAGTCGGAGGCTTAATTACA-3' iNos / Nos2 AF049656 / NM_010927 / NM_012611 Fwd: 5'- CTATCTCCATTCTACTACTACCAGATCGA-3' Rev: 5'- CCTGGGCCTCAGCTTCTCAT-3' Probe: 5'- CCCTGGAAGACCCACATCTGGCAG-3' Ldlr NM_010700 / NM_175762 Fwd: 5'-GCATCAGCTTGGACAAGGTGT-3' Rev: 5'-GGGAACAGCCACCATTGTTG-3' Probe: 5'-CACTCCTTGATGGGCTCATCCGACC-3' Lpl NM_008509 / NM_012598 Fwd: 5'-AAGGTCAGAGCCAAGAGAAGCA-3' Rev: 5'-CCAGAAAAGTGAATCTTGACTTGGT-3' Probe: 5'-CCTGAAGACTCGCTCTCAGATGCCCTACA-3' Lrat NM_023624 Fwd: 5'-TCCATACAGCCTACTGTGGAACA-3' Rev: 5'-CTTCACGGTGTCATAGAACTTCTCA-3' Probe: 5'-ACTGCAGATATGGCTCTCGGATCAGTCC-3' Pck1 NM_011044 / NM_198780 Fwd: 5'-GTGTCATCCGCAAGCTGAAG-3' Rev: 5'-CTTTCGATCCTGGCCACATC-3' Probe: 5'-CAACTGTTGGCTGGCTCTCACTGACCC-3' Ppargc1 α / Pgc1α NM_008904 / NM_031347 Fwd: 5'-GACCCCAGAGTCACCAAATGA-3' Rev: 5'-GGCCTGCAGTTCCAGAGAGT-3' Probe: 5'-CCCCATTTGAGAACAAGACTATTGAGCGAACC-3' Pnpla2 / Atgl NM_025802 / XM_347183 Fwd: 5'-AGCATCTGCCAGTATCTGGTGAT-3' Rev: 5'-CACCTGCTCAGACAGTCTGGAA-3' Probe: 5'-ATGGTCACCCAATTTCCTCTTGGCCC-3' Pnpla3 NM_054088 Fwd: 5'-ATCATGCTGCCCTGCAGTCT-3' Rev: 5'-GCCACTGGATATCATCCTGGAT-3' Probe: 5'-CACCAGCCTGTGGACTGCAGCG-3'RAR-β Assay on demand, Mm01319677_m1 (ThermoFisher)
Raldh1 Assay on demand, Mm00657317_m1 (ThermoFisher)
Raldh2 Assay on demand, Mm00501306_m1 (ThermoFisher)
Raldh3 Assay on demand, Mm00474049_m1 (ThermoFisher)
Raldh4 NM_178713.4 Fwd: 5'-TGGAGCAGTCTCTGGAGGAGTT-3' Rev: 5'- GAAGTTCAGAACAGACCGAGGAA-3' Probe: 5'- AATCTAAAGACCAAGGGAAAACCCTCACGC-3' Rbp4 NM_011255.2 / XM_215285.3 Fwd: 5'-GGTGGGCACTTTCACAGACA-3' Rev: 5'-GATCCAGTGGTCATCGTTTCCT-3' Probe: 5'-CCCCAGTACTTCATCTTGAACTTGGCAGG-3' Scd1 NM_009127.2 Fwd: 5'-ATGCTCCAAGAGATCTCCAGTTCT-3' Rev: 5'-CTTCACCTTCTCTCGTTCATTTCC-3' Probe: 5'-CCACCACCACCATCACTGCACCTC-3' TGF-β1 NM_021578.1 Fwd: 5'-GGGCTACCATGCCAACTTCTG-3' Rev: 5'-GAGGGCAAGGACCTTGCTGTA-3' Probe: 5'-CCTGCCCCTACATTTGGAGCCTGGA-3' TGF-β1 NM_021578.1 Fwd: 5'-GGGCTACCATGCCAACTTCTG-3' Rev: 5'-GAGGGCAAGGACCTTGCTGTA-3' Probe: 5'-CCTGCCCCTACATTTGGAGCCTGGA-3' Timp1 NM_001044384.1 / NM_011593.2 Fwd: 5'-TCTGAGCCCTGCTCAGCAA-3' Rev: 5'-AACAGGGAAACACTGTGCACAC-3' Probe: 5'-CCACAGCCAGCACTATAGGTCTTTGAGAAAGC-3' TNF-α NM_013693 / NM_012675 Fwd: 5'- GTAGCCCACGTCGTAGCAAAC-3' Rev: 5'- AGTTGGTTGTCTTTGAGATCCATG-3' Probe: 5'- CGCTGGCTCAGCCACTCCAGC-3' Ucp2 NM_011671.2 Fwd: 5'-CGAAGCCTACAAGACCATTGC-3' Rev: 5'-ACCAGCTCAGCACAGTTGACA-3' Probe: 5'-CAGAGGCCCCGGATCCCTTCC-3'
